Synthesis of Nanochitosan from Cambarus Bartonii Waste and its Utilization in The Removal of Polycyclic Aromatic Hydrocarbons in Surface Water from Ifite Ogwari, Southeastern Nigeria

Document Type : Research Paper

Authors

1 Department of Pure and Industrial Chemistry, Nnamdi Azikiwe University, Awka, Nigeria

2 Department of Pharmaceutical and Medicinal Chemistry, Nnamdi Azikiwe University, Awka, Nigeria

Abstract

Access to potable water is a global problem, especially in rural communities of the world. Ifite Ogwari is a rural community in Southeastern Nigeria that depends mostly on surface water for its water needs such as drinking. This work explored the use of nanochitosan for removing polycyclic aromatic hydrocarbons (PAHs) contained in the surface water obtained from Ifite Ogwari. Chitosan extracted from Cambarus bartonii waste (CbC) and artificial chitosan (ArC) supplied by ChitoLytic were reduced to nanochitosan particles. The nanochitosan particles were characterized using Dynamic Light Scattering (DLS), also known as Photon Correlation Spectroscopy, Fourier Transformed Infrared Spectroscopy (FTIR), Scanning Electron Microscopy (SEM), and X-ray Diffraction (XRD). PAHs were assessed in untreated and treated water samples using Gas Chromatography-Flame ionization Detector (GC-FID). Results of DLS particle size analyzer ranged between 55.78 nm and 92.64 nm. FTIR gave bands at 2922.2 cm−1 and 1405.2 cm−1 representing C–H and C–N groups, respectively, while the band at 1025.0 cm−1 stretches C–O–C groups. SEM results show interconnected microporosity while XRD results demonstrate sharp peaks at 2θ = 8.20o, 32.08 o, 38.00 o and 45.76 o for nanochitosan from Cambarus bartonii waste (CbNC) and 2θ = 20.21 o, 31.63 o and 45.48 o for nanochitosan from artificial chitosan (ArNC). Nine PAHs were detected in the untreated water while seven were below detectable limit but none was detected in the treated water samples. This study indicates that nannochitosan synthesized from C. bartonii waste could be used for the removal of PAHs from contaminated surface water.

Keywords

Full Text

INTRODUCTION

Water resources around the world could contain a variety of contaminants that, at high concentrations, could increase the risk of contracting a number of diseases in both adults and children, including serious diseases like gastrointestinal illnesses, developmental effects such as cognitive disorders, endocrine disruption, and cancer [1]. The primary sources of drinking water in the majority of rural communities in Nigeria include surface water, such as rivers, lakes, and reservoirs, as well as groundwater aquifers, which are subsurface layers of permeable soil and rock that contain substantial amounts of water. The rural community of Ifite Ogwari in Anambra State is not an exception.

Previous works by Okafor and co-workers indicated that surface and groundwater sources from Ifite Ogwari are polluted by heavy metals and polycyclic aromatic hydrocarbons [2, 3]. Scholars recently have also found PAHs as well as heavy metals in food and food products [4 - 6], beverages [7 – 10], meat and meat products [11, 12] etc.

 Globally, there exist many polycyclic aromatic hydrocarbons, primarily because of persistent anthropogenic pollution sources. As a result of their innate characteristics, including hydrophobicity, thermostability, and heterocyclic aromatic ring structures, PAHs are resistant and extremely persistent in the environment. For a variety of living forms, PAH pollution have been found to be extremely toxic, mutagenic, carcinogenic, teratogenic, and immunotoxicogenic [13-15].

Chitosan, a linear polysaccharide composed of randomly distributed β-(1→4)-linked d-glucosamine and N-acetyl-d-glucosamine has been applied in the treatment of wastewater and water bodies contaminated with heavy metals [16, 17]. Chitosan is also employed in drug manufacturing and numerous other applications such as normalizing blood pressure and cholesterol levels, treating obesity, speeding up the healing of wounds, and as flocculant, coagulant, food additive, weight loss, etc. [18]. The literature abounds with numerous studies on the use of chitosan as a flocculant, coagulant, and as adsorbent for removing contaminants such as heavy metals, dyes, pesticides, antibiotics as well as biological contaminants from wastewater [19 – 21].

The aim of the present study is to adsorb the PAHs in surface water obtained from Ifite Ogwari using synthesized nanochitosan. Previous researches have covered the use of chitosan for wastewater treatment [19 – 21]. Nonetheless, there is little or no existing literature on the extraction of nanochitosan from Cambarus bartonii waste for the treatment of surface water from any Nigerian rural community.

 

EXPERIMENTAL SECTION

Procurement of raw materials

Crayfish waste obtained from Eke-Awka market was made free from dirt before it was pulverized into powder using a dry grinder and sieved with a 2mm sieve. Low molecular weight chitosan (artificial chitosan) was supplied by ChitoLytic from North America. Chemicals and reagents used were potassium hydroxide (KOH), acetic acid (CH3COOH), hydrochloric acid (HCl), sodium hydroxide (NaOH), lead nitrate (Pb(NO3)2), sodium acetate (CH3COONa), sodium tripolyphosphate (Na5P3O10), dihydrogen phosphate (H2PO4), distilled water etc. All reagents used were of analytical grade.

 

Extraction of Chitin and Chitosan

Chitosan was prepared following the method described by Okoya et al. [22] with modifications as adapted by Okafor et al. [23]. The crayfish waste was pulverized into powder using a dry grinder and sieved with 2 mm sieve. 50 g of powdered crayfish waste, with a particle size of less than 2 mm, were weighed into a 500 mL beaker, and 200 mL of 4% (w/v) NaOH was added with constant stirring for 6 h at 80 °C and filtered. The residue was washed with distilled water until it was free of base and then dried at 100 °C for 2 h. The deproteinized crayfish waste residue was then poured into a 250 mL conical flask and 100 mL of 3 % (v/v) 1 mol/dm3 HCl was added and placed on a magnetic stirrer for 3 h at 30°C to demineralize it. The content was filtered and the residue washed until it was free of acid. The acid free residue was then dried at 90 °C for 1 h. A snow-white residue called chitin was obtained. The chitin was poured into a 250 mL conical flask for deacetylation. A 50% (w/v) NaOH solution was added, stirred at 30°C for 4 h and filtered. After filtration, the residue, which is chitosan was washed until the filtrate was neutral and dried at 90°C for 1 h, then stored for further studies.

 

Chitosan Modification

A 1M H2PO4 was added to 100 g of chitosan and allowed to stand for 30 min and after which the liquid was discarded. The wet chitosan was spread on a stainless-steel tray and dried at 50 °C in a forced air oven. After 24 h, the temperature was raised to 180 °C and kept at that temperature for 90 min. The resulting corn-like silks were removed, allowed to cool, and washed in hot deionized water (60- 80 °C) until the liquid did not turn cloudy when mixed with 20 mL lead nitrate solution buffered to pH 4.8 in 0.03 M acetic acid and 0.07 M sodium acetate buffer. This washing step was aimed to remove free phosphate in the corn silk. The wet chitosan was dried at 50 °C for 24h, removed and stored for further examination.

 

Preparation of Nanochitosan

The reduction method used by Okafor et al. [24] was utilized to synthesize nanochitosan. Two grams of chitosan were dissolved in 50 mL of 1% acetic acid with constant stirring, and the pH of the resulting solution was kept at 4 using 1M HCl. Additionally, 100 mg of sodium tripolyphosphate was dissolved in 100 mL of 1M NaOH, and the resulting solution was added to the mixture of acidified chitosan in drops with the help of a syringe until precipitates were fully formed. Using a homogenizer, the mixture was centrifuged at 1000 rpm for five minutes. A filter paper with a 0.2 µm pore size was used to filter the mixture. The nanochitosan was collected as the residue.

 

Characterization of Nanochitosan

Particle Size Analysis

The particle size distribution of the samples was measured by photon correlation spectroscopy (PCS) using a Malvern Zetasizer Nano ZS laser particle size analyzer following a protocol described by Akbari et al. [25] with modifications. The instrument was equipped with a He-Ne laser source wave length (wavelength = 633 nm) and at a scattering angle of 1730o. The dispersion concentration was around 0.1 g/L. The suspension was prepared by dispersing the powder in distilled water and treated for 60 mins in an ultrasonic bath to obtain a well-dispersed suspension.

 

Fourier Transform Infrared Analysis

Buck scientific M530 USA FTIR was used for the analysis. This instrument was equipped with a detector of deuterated triglycine sulphate and beam splitter of potassium bromide. The software of the Gram A1 was utilized to obtain the spectra and to manipulate them. Approximately 1.0 g of the sample was added to 0.5 mL of nujol, mixed properly and placed on a salt pellet. During measurement, FTIR spectra were obtained within the frequency range of 4,000 – 600 cm-1 and co-added at 32 scans and at 4 cm-1 resolutions. FTIR spectra were displayed as transmitter values.

 

Scanning Electron Microscopy (SEM) Analysis

The scanning electron microscopy (SEM) was performed to examine the physical structural changes of samples using SEM model (Phenom ProX, by phenomWorld Einhoven-the Netherlands). The sample was placed on double-sided adhesive which was on a sample stub, coated with a sputter coater by Quorum technologies model (Q150R) with a 5nm layer of gold. Thereafter, it was taken to the chamber of the SEM machine where it was viewed via NaVCaM for focusing and little adjustment. It was then transferred to the SEM mode, focused, and brightness and contrast were automatically adjusted; afterward, the morphologies of different magnifications were stored in a USB stick. After the morphology was acquired, its elemental composition was obtained via the software in the machine.

 

X-ray Diffraction (XRD) Analysis

The sample was prepared and compressed in the flat sample holder to create a flat, smooth surface that was later mounted on the sample stage in the XRD cabinet. The reflection-transmission spinner stage with theta-theta settings was employed to analyze the sample. With a two-theta step of 0.026261 at an average speed of 8.67 seconds per step, the two-theta starting position was 4o and ended at 75o. The tension was 45 VA, and the tube current was set at 40 mA. A Programmable divergent slit with a 5 mm width mask and Gonio Scan were used. The intensity of diffracted X-rays was continuously recorded as the sample and detector rotated through their respective angles. The results were commonly presented as peak positions at 2θ and X-ray counts (intensity).

 

Removal of PAHs contaminated surface water by batch adsorption method

0.2 g of each nanochitosan was weighed in two separate conical flasks. 40 mL of the contaminated water sample was measured and poured into each of the conical flask containing the modified chitosan particles. The orbital shaker was turned on and flasks containing the samples were placed on it to stand for 1hour at room temperature and the speed of 200 rpm. After 1hour, the samples were taken off from the shaker and filtered. The filtrates were taken for further analysis.

 

Extraction and analysis of PAHs in water samples

The method used by Okafor et al. [2] was adopted for the extraction and analysis of the water samples. All water samples were extracted without filtering to reflect exposure levels of PAHs in the water samples.  A separating funnel was used for the extraction of PAHs from the water samples. The extraction of water samples and their spiked duplicates for PAHs followed a common procedure: 250 mL of separating funnel containing 100 mL of an equal mixture of chromatographic grade DCM/n-hexane. The samples were extracted for 2 hours under reflux. The crude extracts were concentrated to a volume of nearly 2 mL using a rotary vacuum evaporator. The resulting concentrates were purified NY short-column silica gel chromatography using DCM as the eluting solvent. The eluates were further reduced to a final volume (2 mL) using nitrogen gas and reconstituted with 2 mL of chromatographic-grade isooctane. The purified extracts (in sealed vials) were kept in the fridge until conducting analysis by GC- FID.

The samples were analyzed with 7890A Agilent Gas chromatograph coupled with a HP5 column (30m x 0.32mm x 0.25um). 1 µL was injected into the GC for the determination of PAH under the following oven condition:

Initial Temp: 60’C, Held for 1min

Ramp rate 1:   Increased to 210’C @ 12’C/min

Ramp rate 2:   Increased to 320’C @ 8’C/min

(Final Temp.) Held 5mins

Total run time:               32.25mins

Detector Temp:             325’C

The concentration of analytes was determined by the peak area of samples against those of the standard with which the equipment was calibrated.

 

RESULTS AND DISCUSSION

Dynamic Light Scattering (DLS) Analysis

The Brownian motion of particles in a dispersion is used in DLS analysis to determine particle size. When a particle suspension is subjected to a monochromatic wave of light, the wavelength of the light changes after striking the particles, and a detector detects the signal.

Four samples were analyzed for particle sizes in order to compare the differences in the sizes of chitosan extracted from Cambarus bartonii (CbC), nanochitosan prepared from CbC (CbNC), low molecular weight artificial chitosan (ArC), and nanochitosan prepared from ArC (ArNC). Table 1 shows the results of the particle sizes of the four products.

The results indicated that there was actually a reduction in size from chitosan to nanochitosan in the extracts. Sample CbC was reduced from 66.55 nm to 55.78 nm (CbNC), while ArC (92.64 nm) was reduced to 68. 82 nm (ArNC). The reduction in the particle sizes of CbC and ArC shows an increase in the surface area of the nanochitosan particles (CbNC and ArNC). The results of DLS in the present work are in good agreement with the results reported by Yusefi et al. [26].

 

Fourier Transform Infrared (FTIR)

FTIR provides information on the chemical structure of the carbon material. FTIR studies were conducted to reveal the probable chemicals contributing to the reduction of nanoparticles. Tables 2 and 3 illustrate the peak wavelengths and functional groups of CbNC and ArNC, respectively. The carbon matrix of organic substances like chitosan does not consist of carbon atoms alone, but is also formed by other hetero-atoms like hydrogen, oxygen, nitrogen, halogens, sulphur, phosphorus, etc. These hetero-atoms are bonded to the edges of the carbon layers which govern the surface chemistry of the adsorbent sample which are detected in both CbNC and ArNC, enabling them to act as an adsorbent. The peaks at 2646.4 cm-1, 2325.9 cm-1 and 2284.9 cm-1 in ArNC indicate the presence of -O-H in phosphorus and oxyacids, and P-H in phosphines, respectively, showing the abundance of heteroatoms. ArNC containing more of these hetero-atoms bonded to their carbon layers as indicated in the FTIR spectra (Figs. 1 and 2) is expected to behave as a better adsorbent compared to CbNC. These results align favorably with those obtained by Divya et al. [27], Rubini et al. [28] and Alshehri et al. [29]. Moreover, the results obtained by Okafor and his coworkers [23, 24] were in agreement with the result of this study.

The bands at 2,922.2 cm−1 and 1405.2 cm−1 represent C–H and C–N groups, respectively, which clearly indicate the absorption spectrum of chitosan [30] and the band at 1025.0 cm−1, stretches C–O–C groups [31]. Furthermore, other researchers have also reported FTIR spectrum for chitosan nanoparticles revealing some variations in the obtained data [32 – 36].

 

Scanning Electron Microscopy (SEM)

SEM analysis demonstrated that the nanochitosan particles had a long thin crystal structure on a non-smooth surface with interconnected granular microporosity (Figs. 1 and 2). This was in accordance with the findings of Mohanasrinivasan et al. [37] that reported a non-homogenous and non-smooth surface structure of nanochitosan in their work. Interconnected granular microporosity was observed more in ArNC favoring its effectiveness as an adsorbent in comparison with CbNC. Chitosan with a long, thin crystal structure on a smooth surface was also reported [38].

 

X-ray Diffraction

X-ray diffraction is known to be a non-destructive technique that works effectively with materials that are wholly or partly crystalline. Figs. 3 and 4 illustrate that CbNC and ArNC have four and three distinct sharp characteristics diffraction peaks, respectively. The peaks at 2θ = 8.20o, 32.08 o, 38.00 o and 45.76 o correspond to CbNC, while those at 2θ = 20.21 o, 31.63 o and 45.48 o are attributed to ArNC.

The sharper peaks provide evidence of a denser crystalline structure [29]. The characteristic peaks of chitosan are reported in the ranges of 2θ= 9.9 - 10.7 and 19.8 - 20.7 [39]. Furthermore, the crystalline nature of shrimp-derived chitosan nanoparticles was determined by the most intense diffraction peaks at values of 11.7 and 20.2° [40]. The difference in the number of diffraction peaks in the two nanochitosan particles indicates that ArNC has denser crystalline structure than CbNC. However, the XRD diffraction patterns obtained by Thamilarasan et al. [41] were located at 2θ = 12.67°, 19.04°, 23.27°, 26.37°, 34.83° and 39.24° which do not align with the findings presented by Zahedi et al. [40]. Nevertheless, these results favorably agree with the findings reported by Okafor et al. [24] who obtained similar results in which diffraction peaks were located between 2θ = 8.1o and 20.1 o for chitosan particles and at 2θ =33.6o, 43.33o and 46.5o for Ag-nanoparticles.

 

PAHs analysis

Three water samples were analyzed for the 16 USEPA priority PAHs namely: untreated water sample (UWS) from Isiachala stream in Ifite Ogwari, South-East Nigeria, water sample from Isiachal stream treated with CbNC (WCbNC), and water sample from Isiachala treated with ArNC (WArNC).

The fingerprint (GC chromatogram) of UWS is represented in Fig. 5, while those of WCbNC and WArNC are illustrated in supplementary Figs. S1 and S2, respectively. It is evident from the fingerprints that 9 PAHs (∑PAHs = 160.9882 ng/ml) were contained in the untreated water sample (UWS) out of the 16 SEPA priority PAHs from Isiachala stream, while none of the PAHs was detected in WCbNC and WArNC as presented in Table 4.

Most of the published literature on the effects of PAHs on human health has mainly focused on their potential to cause cancers of the skin, lung, breast, scrotum, bladder, and colon, etc. [42, 43]. According to Patel et al. [13], PAHs are byproducts of human activity such as forest fires, household and commercial heating appliances, companies and power plants that produce energy and volcanic activity. As a result, they have been detected in water, air, soil, and agricultural products [44]. PAHs in the environment are transferred to these substrates due to direct contact or as a result of precipitation [45].

If inhaled or ingested in high amounts, naphthalene can destroy red blood cells [46]. Flt has also been linked to liver and lung cancers [47], T cell apoptosis [48], a drop in white blood cell counts, and tubular casts in the kidneys [49]. Ugochukwu and Ochonogor reported benzo(a)anthracene’s carcinogenic adverse health effects [50]. For the first time, Tao et al. [51] demonstrated that chrysene exposure damages the liver by activating the aryl hydrocarbon receptor, and that chrysene also protects against oxidative liver damage by activating the nuclear factor erythroid 2-related factor-mediated antioxidant defense mechanism. Benzo[k]fluoranthene (BkF) is known to accelerate the catabolism of 17 estradiol (E2) in human breast cancer cells, according to a study by Arcaro et al. [52]. Women who work in air-polluted BbF environments, such as petrochemical plants and busy roadways, are more likely to have irregular menstrual cycles, aberrant hormone levels, and a greater risk of infertility [53–55]. BbF exposure raises the likelihood of developing cancer [56], causes renal and lung illnesses, and exhibits high reproductive toxicity [57, 58]. Benzo[a]pyrene consumption at high levels during pregnancy has been linked to birth abnormalities and reduced body weight in mouse pups, according to Allamandola [14]. Although it has not been demonstrated that this impact occurs in humans, higher levels of PAH exposure during pregnancy are linked to undesirable birth outcomes, such as low birth weight, premature birth, and heart deformity [59].

High parental exposure is also associated with lower intelligent quotient at age three, increased behavior problems at age six and eight, and childhood asthma. Cord blood of exposed babies show DNA damage that has been linked to cancer [60]. Dibenzo[a,h]anthracene was reported as the most potent carcinogenic [61] as there were no oral or inhalation effects available to assess the carcinogenicity of benzo[ghi]perylene. The result reported for dermal came out negative [62, 63].  It was also reported that benzo[ghi]perylene was administered alongside benzo[a]pyrene to a skin of a mice, and a skin tumor was observed compared to the tumor incidence in mice treated with benzo[a]pyrene alone. This indicates possible carcinogenicity present in benzo[ghi]perylene [64].

The study has demonstrated that nanochitosan particles obtained from C. bartonii could remove all the above PAHs, with the aforementioned health challenges in some rural communities in Nigeria that lack potable water.

 

Proposed mechanism of PAHs adsorption

The mechanism for PAHs adsorption onto artificial and natural nanochitosan could be proposed based on the functional groups as revealed by Infrared analysis. The protonated amino groups from chitosan (NH3+) and the π-electron-rich aromatic structures of the PAHs may interact cationically during the adsorption of PAHs on chitosan-nanoparticles. The adsorption of PAHs may also involve electrostatic interactions and hydrogen bonding due to the presence of amine and hydroxyl group on the adsorbents. The encouraging adsorption results of artificial and natural nanochitosan particles may be attributed to an intramolecular donor–acceptor interaction between the chitosan functional groups and PAHs [65,66].

 

CONCLUSIONS

The data concerning particle sizes reveal that nanochitosan particles from Cambarus bartonii waste and artificial chitosan have lesser sizes than the chitosan extracted from Cambarus bartonii waste and artificial chitosan, respectively. This means that modification was able to decrease the size of the nanochitosan particles. The FTIR analysis shows that carbon matrix of the nanochitosan particles consists of not only carbon atoms, but also other hetero-atoms like hydrogen, oxygen, nitrogen, halogens, sulphur, phosphorus, etc. confirmed by the observed FTIR peaks at 2646.4 cm-1, 2325.9 cm-1, 2284.9 cm-1 etc. Nanochitosan particle from artificial chitosan containing more of these hetero-atoms bonded to their carbon layers as indicated in the FTIR is expected to behave as a better adsorbent compared to the nanochitosan particle from Cambarus bartonii. The significant difference in the morphologies of the nanochitosan particles from Cambarus bartonii and artificial chitosan shows that the interconnected granular microporosity was observed more in the nanochitosan particle from artificial chitosan, favoring its effectiveness as an adsorbent in comparison with nanochitosan particle from Cambarus bartonii waste.

Nanochitosan particles from Cambarus bartonii waste and artificial chitosan have four and three distinct sharp characteristics diffraction peaks, respectively. The peaks at 2θ = 8.20o, 32.08 o, 38.00 o and 45.76 o are related to nanochitosan particle from Cambarus bartonii, waste while those at 2θ = 20.21 o, 31.63 o and 45.48 o correspond to nanochitosan particle from artificial chitosan. The difference in the intensities in the two products indicates that nanochitosan particle from artificial chitosan has denser crystalline structure than nanochitosan particle from Cambarus bartonii waste.

PAHs were present in the surface water sample collected from Isiachala stream in Ifite Ogwari community. This implies that the surface water is not suitable for drinking unless treated. The minimum and maximum concentrations of PAHs in the surface water from Ifite Ogwari were 3.4669 ng/ml (Benzo[a]pyrene) and 82.4078 ng/ml (Chrysene), respectively, and ∑PAHs = 160.9882 ng/ml. The water samples treated with both nanochitosan particles did not indicate the presence of any of the PAHs in them. This implies that the nanochitosan particle from Cambarus bartonii waste could treat PAHs contaminated surface water in the same manner as the artificial chitosan.

In order to reduce exposure to PAHs in the environment, regulatory agencies should regularly monitor and ensure proper water treatment since PAHs are teratogenic, carcinogenic, and mutagenic, and may induce lung, bladder as well as skin cancer. Again, further studies should be carried out on the possibility of using natural raw materials in the extraction of chitosan for use in the treatment of water resources in Nigeria’s rural communities.

 

ACKNOWLEDGMENTS

The authors would like to thank Nnamdi Azikiwe University for supporting the work and the technical staff of Central Laboratory, Nigerian Institute for Oceanography and Marine Research, Victoria Island, Lagos for technical assistance.

 

CONFLICT OF INTEREST

The authors declare no conflicts of interest.

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Nanochemistry Research
Volume 9, Issue 1
January 2024
Pages 42-54

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